Herpes Simplex Virus 1 Infection Activates Poly(ADP-Ribose)
Polymerase and Triggers the Degradation of Poly(ADP-Ribose)
Sarah L. Grady,aJesse Hwang,aLivia Vastag,b* Joshua D. Rabinowitz,band Thomas Shenka
Department of Molecular Biology, Princeton University, Princeton, New Jersey, USA,aand Department of Chemistry and the Lewis-Sigler Institute for Integrative
Genomics, Princeton University, Princeton, New Jersey, USAb
induced drop in NAD?levels required viral DNA replication, was associated with an increase in protein poly(ADP-
of the global human population (36). Like all viruses, HSV-1 de-
pends on the host cell for its replication, and central to this inter-
examined for their dependence and effect on host metabolism,
including cytomegalovirus, Kaposi’s sarcoma-associated herpes-
acid cycle from pyruvate (44). Specifically, inhibition of pyruvate
carboxylase, the enzyme responsible for the conversion of pyru-
vate to oxaloacetate, significantly decreases HSV-1 titers (44). In-
pyrimidine synthesis. An additional, heretofore unexamined,
metabolic alteration during HSV-1 infection is the dramatic de-
crease in the levels of NAD?(44).
NAD?is an important cofactor in many of the reduction-
oxidation (redox) reactions of central carbon metabolism, but it
ribose) polymerase (PARP) superfamily of enzymes as they cata-
lyze the addition of poly(ADP-ribose) (PAR) chains to proteins
(6). PARP-1 is an abundant nuclear enzyme that has been re-
ribosyl)ations (PARylations) in the cell. Of the remaining PARP
(17), and as a consequence, PARP-1 activity has been reported to
have a dominant effect on overall cellular NAD?levels (12).
PARP-1 and PARP-2 (PARP-1/2) are both activated by DNA
damage. The resulting PAR polymers, which can be several hun-
dred units long and are highly negatively charged, help recruit
DNA damage repair machinery to the sites of single- or double-
erpes simplex virus 1 (HSV-1) is an alphaherpesvirus that
strand breaks (3). In the case of significant DNA damage, how-
activity has been implicated in the pathogenesis of several viral
infections. It is necessary for efficient integration of the HIV pro-
viral genome (12) as well as lytic infection by Epstein Barr virus
(26), but its interactions with alphaherpesviruses are largely un-
PARP-1/2 have multiple protein substrates, including many
nuclear enzymes such as DNA polymerases, topoisomerases, and
p53 (25, 33, 38). The acceptors of the majority of PAR chains
automodification inhibits PARP’s catalytic activity, likely by di-
minishing its DNA binding affinity (19, 48). Removal of the PAR
chains occurs via the action of the enzyme poly(ADP-ribose) gly-
cohydrolase (PARG), which possesses both exo- and endoglyco-
chains from protein substrates, and its action on PARP-1/2 effec-
polymerization (7). In humans, PARG is a single gene that codes
for multiple spliced mRNAs. The full-length mRNA produces a
111-kDa (PARG-111) protein that localizes to the nucleus due to
a nuclear localization signal (NLS) present at its N terminus (29).
Received 25 February 2012 Accepted 15 May 2012
Published ahead of print 23 May 2012
Address correspondence to Thomas Shenk, firstname.lastname@example.org.
*Present address: Livia Vastag, Department of Natural Sciences, Castleton State
College, Castleton, Vermont, USA.
Copyright © 2012, American Society for Microbiology. All Rights Reserved.
August 2012 Volume 86 Number 15Journal of Virologyp. 8259–8268jvi.asm.org
Isoforms of 102 and 99 kDa (PARG-102 and PARG-99, respec-
tively) are found in the cytoplasm but have been shown to shuttle
to sites of DNA damage in the nucleus after microirradiation and
gamma irradiation (3, 14, 30). Smaller PARG isoforms of low
abundance are enriched in mitochondria and do not appear to
alter their localization patterns (30, 47).
In this study, we show that HSV-1 replication activates PARP-
PARylation levels. Neither a further decrease nor a rescue of
NAD?levels altered the metabolic effects of HSV-1 on the host
cell, suggesting that, to a large extent, HSV-1 infection supersedes
other cellular signals to control overall metabolic status. The in-
this process. We also found that HSV-1 infection triggered the
proteasome-dependent degradation of the 111-kDa isoform of
HSV-1-infected cells due to the activation of PARP-1/2 and that
infection actively modulates the balance between PARP and
MATERIALS AND METHODS
Cells, viruses, and reagents. Primary human fibroblasts were used be-
tween passages 8 to 15. Cells were grown in Dulbecco’s modified Eagle
medium (DMEM) with 4.5g/liter glucose (Sigma) supplemented with
10% fetal bovine serum, and 100 ?g/ml penicillin and streptomycin (In-
vitrogen). HSV-1 strain F (9) and a C116G/C156A mutant (22) were
kindly provided by B. Roizman (University of Chicago) and grown in
Vero cells. Stocks were produced by pooling cell-associated virus, ob-
infectious focus assay and are expressed as infectious units (IU). Briefly,
postinfection (hpi) with methanol at ?20°C. Foci were identified using a
(39) and goat anti-mouse Alexa Fluor 488-conjugated secondary anti-
(DMSO) used at 20 nM. Olaparib (LC Laboratories) was dissolved in
in DMEM to a stock concentration of 6 mM, filter sterilized (0.22-?m
pore size), and used at 3 mM. Acyclovir (Sigma) was dissolved in DMSO
and used at 1 ?M. MG132 (Cayman Chemicals) was dissolved in DMSO
and used at concentrations of 200 nM to 10 ?M. Phosphonoacetic acid
(Sigma) was dissolved in ethanol and used at 400 ?g/ml.
Metabolic analysis. Fibroblasts were grown to confluence and main-
tained in the presence of serum for 3 to 5 days. Cells were then washed
with the equivalent volume of virus-free DMEM. After a 1-h adsorption
period, cells were washed, and fresh medium was added. Following vari-
ous time intervals, the medium was aspirated from cells and an 80:20
olism. Metabolites were then extracted as described previously (49). Ex-
tracts were dried under nitrogen gas and resuspended in high-perfor-
mance liquid chromatography (HPLC)-grade water. Samples were
centrifuged at 15,000 ? g for 5 min to remove any remaining particulate
To quantify the levels of metabolites in extracts, we utilized an untar-
geted analysis approach using liquid chromatography (LC) coupled to a
stand-alone Orbitrap mass spectrometer (Thermo Fisher Scientific Exac-
a mass resolution of 100,000 (24). Compound identification is based on
retention time on the LC column, and compound mass was measured to
within an accuracy of 2 ppm. Peaks were identified and metabolites were
quantified with the Metabolomic Analysis and Visualization Engine
(MAVEN) software package (27).
For each time point in experiments comparing infected and unin-
fected cells, an additional plate for each treatment was processed for
packed cell volume measurements. Briefly, the cells of one 35-mm plate
was centrifuged at 2,000 ? g for 5 min before a reading was taken (43).
Packed cell volume measurements were used to normalize the metabolite
levels between samples. All metabolite levels are averages of duplicate
Protein analysis. For Western blot assays, fibroblasts were grown to
adsorption period, cells were washed once, and fresh medium was added.
At the indicated times postinfection, cells were washed with phosphate-
cell lysates, cells were lysed in radioimmunoprecipitation assay (RIPA)
light buffer (50 mM Tris-HCl, pH 8.0, 1% NP-40, 0.1% SDS, 150 mM
NaCl, 0.1% Triton X-100, 5 mM EDTA) with protease inhibitors (Roche
Applied Science). Protein concentrations were determined by Bradford
assay (Bio-Rad). Proteins were separated by electrophoresis in an 8% or
membranes. Membranes were blocked in PBS–0.1% Tween (PBST) with
5% nonfat dry milk (NFDM). All antibodies were diluted in PBST–5%
NFDM. Mouse monoclonal antibodies used in this study included anti-
PARG (MABS61; Millipore) (1:1,000), anti-PAR (1020; Tulip Biosci-
hybridoma supernatant), anti-?-tubulin (T6199; Sigma) (1:5,000), and
horseradish peroxidase (HRP)-conjugated anti-?-actin (49900; Abcam)
(1:20,000). The rabbit polyclonal antibody used was anti-PARP-1 (9542;
Cell Signaling) (1:1000), and the rabbit monoclonal antibody used was
anti-phospho-H2AX (9718; Cell Signaling) (1:1,000). The rabbit antise-
rum used was directed against the N-terminal domain of PARG (29) and
was a kind gift of Myron Jacobson (University of North Texas) (1:5,000).
The goat polyclonal antibody used was anti-promyelocytic leukemia
with PBST and subsequently probed with goat anti-rabbit, goat anti-
mouse, or mouse anti-goat HRP-coupled secondary antibodies (1:5,000;
Jackson ImmunoResearch). Proteins were visualized by chemilumines-
cence using an ECL detection system (Amersham).
For immunofluorescence assays, fibroblasts on glass coverslips were
grown to confluence and serum starved for 24 h before infection. At in-
dicated times, cells were washed in PBS and fixed in methanol/acetone
Primary antibodies were diluted in PBS supplemented with 10% human
were washed three times with PBS–0.2% Tween between primary and
secondary antibody incubations. Secondary antibodies were diluted in
PBS–10% goat serum and incubated as described for primary antibodies.
Mouse monoclonal antibodies used for indirect immunofluorescence
kind gift of R. Everett, University of Glasgow) (1:3,000) and anti-phos-
secondary antibodies conjugated to Alexa Fluor 488 or 546 (Invitrogen)
(Invitrogen). Confocal images were obtained using Zeiss LSM510 laser
Grady et al.
jvi.asm.orgJournal of Virology
Transfections. For siRNA transfections, fibroblasts were grown to
80% confluence in 24-well dishes and maintained in growth medium
without antibiotics. Cells were transfected with 10 pmol of siRNA
using Lipofectamine RNAiMAX (Invitrogen) following the manufac-
turer’s instructions. For experiments using HSV-1, transfected cells
were allowed to incubate for 3 days before being infected with HSV-1
at a multiplicity of 3 IU/cell. Medium and cells were harvested at 18 to
24 hpi as described above. Double-stranded siRNAs used for this ex-
periment were purchased from Sigma and included PARP-1
(Hs02_00332177), PARG (Hs01_00128017), and a universal nontar-
geting control (SIC0001).
For plasmid transfections, fibroblasts were grown to 70 to 80% con-
fluence and washed with Opti-MEM (Invitrogen) before transfection
versity of Chicago), using Lipofectamine LTX with Plus reagent (Invitro-
gen) according to the manufacturer’s instructions. When MG132 was
used, drug was added 1 h after transfection in Opti-MEM. Cells were
supplemented with growth medium lacking antibiotics 4 h after transfec-
tion and were allowed to grow for an additional 24 h. Samples were col-
lected for Western blot assays as described above.
Statistical analysis. Data are expressed as averages ? standard devia-
tions (SD). Statistical analysis was done using Student’s t test, and signif-
icance was set at a P value of ?0.05.
HSV-1 infection (44). To confirm this observation, we initially
infection of human fibroblasts. Confluent cells were serum
starved for 24 h and then infected or mock infected. Serum star-
vation synchronizes cells in G0and decreases variability in the
infection environment between cells (4). This treatment also re-
moves extraneous metabolites that would confound mass spec-
FIG 1 HSV-1 infection of human fibroblasts leads to the depletion of cellular NAD?pools. (A) NAD?levels over the course of HSV-1 infection (3 IU/cell) of
serum-starved fibroblasts, normalized to packed cell volume. Values are averages of duplicate biological experiments (?1 SD). (B) NAD?levels after pretreat-
ment with 20 nM FK866, an inhibitor of NAD?biosynthesis, or DMSO. Cells were treated with FK866 for 24 h before being washed and infected with HSV (3
cells. Each treatment is the average of triplicate biological experiments (?1 SD). (D) Metabolite abundances after treatment with 20 nM FK866 as described
above. Metabolite concentrations are expressed relative to mock-treated cells, and all ratios were log2transformed. AU, arbitrary units. *, P ? 0.05.
Activation of PARP by HSV-1
August 2012 Volume 86 Number 15jvi.asm.org 8261
in infection to the time of maximum virus output at approxi-
mately 24 h postinfection (hpi), and NAD?was measured by liq-
uid chromatography-high resolution
(LC-MS). HSV-1 infection triggered a dramatic, time-dependent
decrease in NAD?levels, such that infected cells had only ?20%
of the NAD?levels seen in uninfected cells by 24 hpi (Fig. 1A).
Levels of NADH also decreased during infection, while overall
NAD?/NADH ratios stayed relatively stable. This virus-induced
decrease in NAD?levels could have been due to increased deple-
tion of cellular NAD?pools or inhibition of NAD?biosynthesis.
To distinguish between these possibilities, we pretreated fibro-
blasts with 20 nM FK866, a specific and noncompetitive inhibitor
of nicotinamide phosphoribosyltransferase (NAMPT), a key en-
zyme in the salvage-based synthesis of NAD?from nicotinamide
(15). After 24 h of FK866 treatment, uninfected fibroblasts had
NAD?levels depleted by more than 50%. Following this 24-h
metabolites were analyzed at 18 hpi. HSV-1 caused a drop in
FIG 2 Depletion of NAD?during HSV-1 infection is due to PARP-1/2 activation. (A) Analysis of poly(ADP-ribosyl)ation over the course of infection.
H2O2at 37°C served as a positive control for PARP activation. Whole-cell lysates were analyzed by Western blotting with an antibody specific to poly(ADP)-
ribose. ?-Actin was used as a loading control. (B) Analysis of PARP-1 protein levels over the course of infection with HSV-1. Serum-starved fibroblasts were
were harvested, and PARP-1 levels were monitored by Western blotting. A caret (?) indicates a 30-min incubation with 10 mM sorbitol, a positive control for
IU/cell) or mock infected, and drug was applied at 1 hpi. Metabolites were extracted at 18 hpi, and NAD?levels were normalized to packed cell volume. Values
are averages of duplicate experiments (?1 SD). (D) Production of infectious HSV-1 virions in cells treated with 50 nM Olaparib as described above. Values are
representative of virus yield at 18 hpi and are expressed relative to DMSO-treated cells (?1 SD). (E) Analysis of poly(ADP-ribosyl)ation after PARP-1/2
inhibition. Serum-starved fibroblasts were infected with HSV-1 (3 IU/cell) or mock infected. At 1 hpi, 50 nM Olaparib was applied, and cells were harvested at
18 hpi. H2O2-treated cells were pretreated with 50 nM Olaparib for 1 h before H2O2application. Whole-cell lysates were analyzed by Western blotting. (F)
Metabolite abundances after treatment with 50 nM Olaparib as described above. Metabolite concentrations are expressed relative to equivalent mock-treated
cells, and all ratios were log2transformed. AU, arbitrary units; M, mock infected. *, P ? 0.05.
Grady et al.
jvi.asm.org Journal of Virology
NAD?levels in FK866-treated cells that was significantly greater
time (Fig. 1B). This argues that the virus-induced depletion of
NAD?was due to increased consumption of existing pools. This
the drug caused no significant change in viral titers (Fig. 1C).
In addition to its effects on NAD?, HSV-1 infection of serum-
starved fibroblasts also triggers the accumulation of metabolites
from upper glycolysis and several intermediates in the synthesis
pathways of nucleotides (44). FK866 pretreatment of uninfected
a much weaker extent than seen after HSV-1 infection (Fig. 1D).
When FK866-treated cells were subsequently infected, the meta-
bolic results were similar in magnitude to the changes seen with
HSV-1 infection alone.
The depletion of NAD?levels during infection results from
PARP activation. NAD?is an essential cofactor in many of the
redox reactions of central carbon metabolism, during which
NAD?is converted to and from its reduced form, NADH. Unlike
NAD?as a substrate toward the formation of PAR chains. As the
primary consumers of NAD?, enzymes of the PARP superfamily,
and especially PARP-1, are largely responsible for overall cellular
NAD?levels (6). Given the decrease in NAD?levels seen during
1/2, triggering NAD?consumption and increasing PAR chain
synthesis. We found that over the course of HSV-1 infection, and
especially at early time points, the abundance of PARylated pro-
teins increased (Fig. 2A). This was not due to an increase in total
PARP-1 or its apoptosis-specific 85-kDa cleavage product (40),
which remained constant throughout infection (Fig. 2B). To fur-
ther test for a role of PARP-1/2, we measured NAD?in both
infected and mock-infected cells after treatment with Olaparib,
which specifically and potently inhibits both enzymes (23, 28).
Whereas Olaparib had no effect on NAD?levels in uninfected
cells, it completely restored the NAD?levels of infected cells (Fig.
as drug treatment did not significantly change viral titers (Fig.
2D). The NAD?rescue phenotype was also recapitulated in the
matically after Olaparib treatment (Fig. 2E). In a control experi-
ment, when fibroblasts were pretreated with Olaparib before ex-
posure to hydrogen peroxide, PARylated protein levels also
decreased, consistent with previous data showing that H2O2acti-
vates PARP activity (45, 48). PARP-1/2 inhibition had little effect
on the metabolic status of either uninfected or infected cells (Fig.
nals in determining the overall metabolic state of the cell.
NAD?levels in infected cells were also rescued after applica-
HSV-1 infected cells. (A) NAD?levels after treatment with 3-aminobenz-
amide (ABA). Serum-starved fibroblasts were infected (3 IU/cell) or mock
infected, and 3 mM ABA or DMEM was applied at 1 hpi. Metabolites were
extracted at 24 hpi, and NAD?levels were normalized to packed cell volume.
Values are averages of duplicate experiments (?1 SD). (B) Production of
infectious HSV-1 virions in cells treated with 3 mM ABA or DMEM as de-
scribed above. Values are representative of virus yield at 18 hpi and are ex-
pressed relative to mock-treated cells (?1 SD).
FIG 4 Viral DNA replication is required for PARP activation. (A) NAD?
replication, or DMSO. Serum-starved fibroblasts were infected (3 IU/cell) or
hpi, and NAD?levels were normalized to packed cell volume. Values are
averages of duplicate experiments (?1 SD). (B) Production of infectious
HSV-1 virions in cells treated with 1 ?M acyclovir as described above. Values
are representative of virus yield at 18 hpi and are expressed relative to DMSO-
treated cells (?1 SD). (C) Analysis of poly(ADP-ribosyl)ation after inhibition
of HSV-1 DNA replication. Serum-starved fibroblasts were infected with
cells were harvested at 18 hpi. H2O2-treated cells were pretreated with 1 ?M
acyclovir for 1 h before H2O2application. Whole-cell lysates were analyzed by
Activation of PARP by HSV-1
August 2012 Volume 86 Number 15jvi.asm.org 8263
tion of 3 mM 3-aminobenzamide (Fig. 3A), a broad-spectrum
inhibitor of ADP-ribosylation (42). This drug had a modest
ascertained whether this reduction resulted from inhibition of
PARP-1/2 or from a general decrease in ADP-ribosylation events.
A previous report has shown that inhibition of the ADP-ribosyla-
tion activities of tankyrase-1 and tankyrase-2, which would be
altered by 3-aminobenzamide but not Olaparib, reduces HSV-1
We conclude that PARP-1/2 activity is strongly activated in
HSV-1-infected cells, resulting in the consumption of NAD?and
generating PARylated proteins.
Viral DNA replication is required for PARP activation.
PARP-1/2 activation occurs following DNA damage, and it has
been well documented that HSV-1 infection activates certain
that the replication and/or resolution of viral genome concatem-
replication is required for PARP-1/2 activation, we treated cells
with acyclovir, a nucleotide analog that is phosphorylated by the
minating elongation (10). Acyclovir (1 ?M) decreased viral yield
by ?200-fold and completely restored NAD?levels in infected
cells (Fig. 4A and B). Acyclovir also reduced the levels of
PARylated proteins in infected cells, while having no effect on
PARylation following H2O2treatment (Fig. 4C). Thus, we can
conclude that viral DNA replication is necessary for activation of
PARP-1/2 and depletion of NAD?during HSV-1 infection.
To further examine the relationship between viral DNA repli-
cation and PARP-1/2 activation, we measured the phosphoryla-
phorylated on Ser 139 (?H2AX) is a marker of DNA breaks and
increases with increased levels of DNA damage (35). As expected,
total ?H2AX levels increased as HSV-1 infection progressed (Fig.
5A). When infected cells were treated with 400 ?g/ml phospho-
noacetic acid (PAA), which directly inhibits the viral DNA poly-
merase (16) at concentrations sufficient to decrease the viral yield
by ?200-fold, ?H2AX levels were markedly decreased at both 8
and 18 hpi (Fig. 5B to D). PAA also decreased total PARylated
proteins in infected cells while having no effect on PARylation
after application of H2O2(Fig. 5E). Thus, two drugs that inhibit
HSV-1 DNA replication by different mechanisms were both able
to reduce infection-induced PARylation, supporting the hypoth-
esis that viral DNA replication is necessary for PARP-1/2 activa-
active viral DNA replication (Fig. 5F).
FIG 5 HSV-1 DNA replication triggers the accumulation of DNA breaks. (A) Images showing the phosphorylation levels of histone H2AX (?H2AX) over the
course of HSV-1 infection. Serum-starved fibroblasts were infected (1 IU/cell) and fixed at various times. Protein levels and localization were examined by
immunofluorescence using antibodies specific to ?H2AX and HSV-1 ICP4. (B and C) Analysis of H2AX phosphorylation after inhibition of the viral DNA
in cells treated with 400 ?g/ml PAA as described above. Values are representative of virus yield at 18 hpi and are expressed relative to mock-treated cells. (E)
and treated with PAA as described above. H2O2-treated cells were pretreated with PAA for 1 h before H2O2application. Whole-cell lysates were analyzed by
Grady et al.
jvi.asm.org Journal of Virology
ICP0 mediates the proteasome-dependent degradation of
self-inhibited by automodification (6). The enzyme poly(ADP-
PARylated proteins, including PARP-1/2, thus reactivating the
determine if HSV-1 infection alters the balance between PARP
at molecular masses matching the 111- and 102/99-kDa isoforms
band into the 102- and 99-kDa isoforms (data not shown). Upon
HSV-1 infection, the levels of the higher-molecular-mass PARG
decreased; a similar decrease was also seen upon hydrogen perox-
ide treatment. To ensure that all the present bands truly repre-
sented PARG, an siRNA transfection was done using an siRNA
that targeted all isoforms. Both the higher- and lower-molecular-
no change in cell viability, as measured by propidium iodide (PI)
uptake, was seen after transfection (nontargeting siRNA, 8.4% ?
1.2% PI positive; PARG siRNA, 8.7% ? 3.2% positive). To deter-
mine if the higher-molecular-mass PARG band was the 111-kDa
isoform, we used antiserum directed against the N terminus of
PARG, which is spliced out in all the smaller isoforms. The single
treatment as well as with infection (Fig. 6B). Knockdown of all
PARG isoforms by siRNA treatment resulted in a modest, but
infection that occurs when all PARG isoforms are knocked down,
viral gene products were probed with and without PARG siRNA
treatment (Fig. 6D). The levels of VP16, ICP0, and gM remained
the same regardless of siRNA treatment, suggesting that the block
to infection occurs in a gene not tested or at a step after leaky late
ICP0 is an immediate-early viral protein with a RING finger
domain possessing E3 ubiquitin ligase activity that has been
shown to sponsor the proteasome-dependent degradation of a
wide range of cellular proteins, including DNA-dependent pro-
tein kinase (DNA-PK), PML, and centromere protein B
(CENP-B) (13, 34). To determine if the loss of the 111-kDa iso-
form of PARG during HSV-1 infection was a result of protea-
some-dependent degradation, we treated infected cells with
MG132, an inhibitor of the 26S proteasome subunit (11) (Fig.
PML and the 111-kDa PARG isoform at 12 hpi. To test whether
ICP0 was sufficient for PARG-111 degradation, we transfected
fibroblasts with an ICP0 expression vector and probed for PARG
protein levels. One day after transfection, ICP0-expressing cells
vector (Fig. 7B), and this decrease was blocked by MG132 treat-
To confirm that ICP0 was necessary for the degradation of
infection with wild-type virus versus infection with C116G/
C116G/C156A mutant did not induce the degradation of PARG-
111, even at late times when ICP0 expression was high (Fig. 7C).
of ICP0 is responsible for the decrease in PARG-111 levels seen
Interactions between herpesviruses and their host cells, including
changes in metabolic status upon infection, are varied and com-
FIG 6 Levels of a high-molecular-mass PARG isoform decrease during HSV-1 infection. (A) Analysis of PARG protein content in cells transfected with siRNA
directed against PARG. Subconfluent fibroblasts were transfected with 10 pmol of siRNA. Three days later, cells were infected with HSV-1 (3 IU/cell), mock
Subconfluent fibroblasts were transfected with nontargeting (NT) or PARG (P) siRNA as above. Whole-cell lysates were collected 3 days after transfection.
lysates were analyzed by Western blotting. M, mock infected. *, P ? 0.05.
Activation of PARP by HSV-1
August 2012 Volume 86 Number 15 jvi.asm.org 8265
plex. NAD?plays a significant role as a cofactor in multiple reac-
overall energy state of the cell (21). Here, we show that HSV-1
infection dramatically decreases the levels of NAD?in the cell
(Fig. 1) and increases protein PARylation (Fig. 2A). Treatment
ary to PARP-1/2 activation (Fig. 2C to F) and dependent on viral
DNA replication (Fig. 4 and 5E). The levels of ?H2AX increase
during infection but are reduced when the viral DNA polymerase
is inhibited (Fig. 5A to C), which suggests that inhibition of viral
genome replication decreases at least some types of DNA damage
typically induced by infection. As PARP-1/2 is activated by DNA
be due to virus-induced DNA damage activating PARP activity.
the activity of these proteins is critical as removal of PAR auto-
modifications from PARP-1/2 can restore its catalytic activity (7).
Fibroblasts expressed the 111-, 102-, and 99-kDa isoforms of
PARG, and their relative abundances agreed with endogenous
PARG levels previously seen in HeLa cells (29). Knockdown of
all PARG isoforms by siRNA (Fig. 6A) resulted in moderately
decreased HSV titers (Fig. 6B), suggesting that some PARG activ-
ity facilitates HSV-1 replication. It was also shown that the viral
protein ICP0 directed degradation of PARG-111 but not its
smaller isoforms (Fig. 7).
As the relative roles of PARP-1/2 and PARG must be tightly
regulated for the proper response to DNA damage (50) and as
ponents of damage repair (46), it is intriguing that ICP0 mediates
the degradation of the 111-kDa isoform but not the 102- and
99-kDa isoforms of PARG. The differences between these iso-
forms are not well characterized although their overexpression in
HEK293 cells suggests that PARG-111 is nuclear whereas PARG-
102 and -99 are located in the cytoplasm (29). It has also been
suggested that these smaller isoforms may shuttle to the nucleus
lethal (18), and PARG siRNA treatment reducing all PARG iso-
forms lowered virus yields (Fig. 6B), raising the possibility that
selective degradation of PARG-111 allows for optimal viral
growth. Selective PARG-111 degradation in mouse astrocytes has
been shown to slow the rate of nuclear PAR degradation and pro-
tect against PARP-dependent cell death, which is termed
parthanatos (5). In this pathway, PARP-generated PAR polymers
translocate from the nucleus to the cytoplasm, where they bind
cytoplasm, and it eventually enters the nucleus, where it induces
cell death. Thus, ICP0-mediated degradation of PARG-111 could
represent a viral strategy to protect infected cells from premature
death due to excessive production of PAR polymer by preventing
the removal of inhibitory PAR chains from PARP, thereby reduc-
ing its activity at nuclear replication compartments. Retention of
mic PARG in mouse neurons has been shown to decrease cyto-
plasmic PAR levels and inhibit the release of AIF from the mito-
chondria after parthanatos-inducing treatments (1). It remains
unclear whether PARP-1/2 activity contributes positively to the
is PARylated but to unknown effect (2).
the related betaherpesvirus human cytomegalovirus (HCMV)
(44). It is interesting that HCMV does not trigger the loss of
NAD?. The substantial difference in infection kinetics between
these viruses, with HSV-1 replicating its genome earlier after in-
fection and completing its replication cycle much faster than
HCMV, may be the reason for this contrast. Maintaining NAD?
levels would be especially important to HCMV as infection trig-
gers increased flux through glycolysis, which requires NAD?as a
Drug-induced depletion of NAD?levels of uninfected cells by
FK866 gave a metabolic output similar to that seen with infection
infection with Olaparib did not modify the global metabolic state
induced by infection. So while the metabolic alterations induced
FIG 7 The HSV-1 protein ICP0 mediates the proteasome-dependent degra-
dation of a high-molecular-mass isoform of PARG. (A) Analysis of PARG
protein levels after treatment with the proteasome inhibitor MG132. Serum-
starved fibroblasts were infected with HSV (3 IU/cell) or mock infected. Cells
were washed and treated with DMSO or 10 ?M MG132 at 1 hpi. Lysates were
positive control for HSV-induced, proteasome-dependent degradation of a
host cell protein. (B) Subconfluent fibroblasts were transfected with MTS1-
ICP0 or the corresponding empty vector MTS1. Then, 200 nM MG132 or
Analysis of PARG protein levels in the C116G/C156A mutant, which contains
a mutation in the RING finger E3 ubiquitin ligase domain of ICP0. Serum-
starved fibroblasts were infected with HSV F strain or the C116G/C156A mu-
Grady et al.
jvi.asm.orgJournal of Virology
is depleted in uninfected cells, the virus-induced changes are not
themselves dependent on the depletion of NAD?during infec-
tion. Thus, despite the large number of metabolic reactions that
utilize NAD?as a cofactor, HSV-1 infection appears to have a
more dominant role than NAD?levels in determining the overall
metabolic status of the cell.
In summary, we have found that HSV-1 infection activates
This change, however, is not responsible for the other metabolic
effects of infection, nor does it significantly impact viral replica-
tion. By late in infection, PARG-111 is degraded in an ICP0-de-
is inhibitory to the virus. This work provides an explanation for
the profound changes in NAD?levels that occur subsequent to
HSV-1 infection, and it demonstrates that HSV-1 infection ac-
tively alters the fine-tuned balance between PARP and PARG in
vided by B. Roizman, L. Terry, and E. O’Keefe.
This work was supported by grants from the National Institutes of
Health to T.S. (CA82396) and J.D.R. (AI078063). S.L.G. is supported
by a National Science Foundation Graduate Research Fellowship
1. Andrabi SA, et al. 2006. Poly(ADP-ribose) (PAR) polymer is a death
signal. Proc. Natl. Acad. Sci. U. S. A. 103:18308–18313.
2. Blaho JA, et al. 1992. Differences in the poly(ADP-ribosyl)ation patterns
of ICP4, the herpes simplex virus major regulatory protein, in infected
cells and in isolated nuclei. J. Virol. 66:6398–6407.
3. Blenn C, Wyrsch P, Althaus FR. 2011. The ups and downs of tannins as
4. Browne E, Wing B, Coleman D, Shenk T. 2001. Altered cellular mRNA
levels in human cytomegalovirus-infected fibroblasts: viral block to the
accumulation of antiviral mRNAs. J. Virol. 75:12319–12330.
5. Burns DM, Ying W, Kauppinen TM, Zhu K, Swanson RA. 2009.
Selective down-regulation of nuclear poly(ADP-ribose) glycohydrolase.
PLoS One 4:e4896. doi:10.1371/journal.pone.0004896.
6. D’Amours D, Desnoyers S, D’Silva I, Poirier GG. 1999. Poly(ADP-
ribosyl)ation reactions in the regulation of nuclear functions. Biochem. J.
7. Davidovic L, Vodenicharov M, Affar EB, Poirier GG. 2001. Importance
of poly(ADP-ribose) glycohydrolase in the control of poly(ADP-ribose)
metabolism. Exp. Cell Res. 268:7–13.
8. Delgado T, et al. 2010. Induction of the Warburg effect by Kaposi’s
sarcoma herpesvirus is required for the maintenance of latently infected
endothelial cells. Proc. Natl. Acad. Sci. U. S. A. 107:10696–10701.
9. Ejercito PM, Kieff ED, Roizman B. 1968. Characterization of herpes
simplex virus strains differing in their effects on social behaviour of in-
fected cells. J. Gen. Virol. 2:357–364.
10. Elion GB. 1982. Mechanism of action and selectivity of acyclovir. Am. J.
11. Giuliano M, D’Anneo A, De Blasio A, Vento R, Tesoriere G. 2003.
Apoptosis meets proteasome, an invaluable therapeutic target of antican-
cer drugs. Ital. J. Biochem. 52:112–121.
12. Ha HC, et al. 2001. Poly(ADP-ribose) polymerase-1 is required for effi-
cient HIV-1 integration. Proc. Natl. Acad. Sci. U. S. A. 98:3364–3368.
13. Hagglund R, Roizman B. 2003. Herpes simplex virus 1 mutant in which
the ICP0 HUL-1 E3 ubiquitin ligase site is disrupted stabilizes cdc34 but
degrades D-type cyclins and exhibits diminished neurotoxicity. J. Virol.
14. Haince JF, Ouellet ME, McDonald D, Hendzel MJ, Poirier GG. 2006.
radiation-induced DNA damage. Biochim. Biophys. Acta 1763:226–237.
15. Hasmann M, Schemainda I. 2003. FK866, a highly specific noncompet-
itive inhibitor of nicotinamide phosphoribosyltransferase, represents a
novel mechanism for induction of tumor cell apoptosis. Cancer Res. 63:
16. Honess RW, Watson DH. 1977. Herpes simplex virus resistance and
sensitivity to phosphonoacetic acid. J. Virol. 21:584–600.
17. Huber A, Bai P, de Murcia JM, de Murcia G. 2004. PARP-1, PARP-2 and
ATM in the DNA damage response: functional synergy in mouse devel-
opment. DNA Repair (Amst.) 3:1103–1108.
18. Koh DW, et al. 2004. Failure to degrade poly(ADP-ribose) causes in-
creased sensitivity to cytotoxicity and early embryonic lethality. Proc.
Natl. Acad. Sci. U. S. A. 101:17699–17704.
20. Li Z, et al. 2012. Herpes simplex virus requires PARP activity for efficient
replication and induces ERK-dependent phosphorylation and ICP0-
dependent nuclear localization of tankyrase 1. J. Virol. 86:492–503.
21. Lin SJ, Guarente L. 2003. Nicotinamide adenine dinucleotide, a meta-
bolic regulator of transcription, longevity and disease. Curr. Opin. Cell
22. Lium EK, Silverstein S. 1997. Mutational analysis of the herpes simplex
the expression of the essential ?27 gene. J. Virol. 71:8602–8614.
23. Loh VM, Jr, et al. 2005. Phthalazinones. Part 1: the design and synthesis
of a novel series of potent inhibitors of poly(ADP-ribose)polymerase.
Bioorg. Med. Chem. Lett. 15:2235–2238.
24. Lu W, et al. 2010. Metabolomic analysis via reversed-phase ion-pairing
liquid chromatography coupled to a stand alone Orbitrap mass spectro-
meter. Anal. Chem. 82:3212–3221.
25. Malanga M, Pleschke JM, Kleczkowska HE, Althaus FR. 1998. Poly-
functions. J. Biol. Chem. 273:11839–11843.
26. Mattiussi S, et al. 2007. Inhibition of Poly(ADP-ribose)polymerase im-
27. Melamud E, Vastag L, Rabinowitz JD. 2010. Metabolomic analysis and
visualization engine for LC-MS data. Anal. Chem. 82:9818–9826.
28. Menear KA, et al. 2008. 4-[3-(4-cyclopropanecarbonylpiperazine-1-
29. Meyer-Ficca ML, Meyer RG, Coyle DL, Jacobson EL, Jacobson MK.
splice variants yielding isoforms that localize to different cell compart-
ments. Exp. Cell Res. 297:521–532.
30. Mortusewicz O, Fouquerel E, Ame JC, Leonhardt H, Schreiber V. 2011.
PARG is recruited to DNA damage sites through poly(ADP-ribose)- and
PCNA-dependent mechanisms. Nucleic Acids Res. 39:5045–5056.
31. Munger J, Bajad SU, Coller HA, Shenk T, Rabinowitz JD. 2006. Dy-
namics of the cellular metabolome during human cytomegalovirus infec-
tion. PLoS Pathog. 2:e132. doi:10.1371/journal.ppat.0020132.
32. Munger J, et al. 2008. Systems-level metabolic flux profiling identifies
fatty acid synthesis as a target for antiviral therapy. Nat. Biotechnol. 26:
vitro. Biochem. Biophys. Res. Commun. 140:666–673.
34. Paulus C, Nitzsche A, Nevels M. 2010. Chromatinisation of herpesvirus
genomes. Rev. Med. Virol. 20:34–50.
35. Redon CE, et al. 2010. Histone ?H2AX and poly(ADP-ribose) as clinical
pharmacodynamic biomarkers. Clin. Cancer Res. 16:4532–4542.
36. Roizman B, Knipe DM, Whitley RJ. 2007. Herpes simplex viruses, p
2501–2601. In Knipe DM, Howley PM (ed), Fields virology, 5th ed. Lip-
pincott Williams and Wilkins, Philadelphia, PA.
37. Rouleau M, Patel A, Hendzel MJ, Kaufmann SH, Poirier GG. 2010.
PARP inhibition: PARP1 and beyond. Nat. Rev. Cancer. 10:293–301.
38. Satoh MS, Lindahl T. 1992. Role of poly(ADP-ribose) formation in DNA
repair. Nature 356:356–358.
39. Showalter SD, Zweig M, Hampar B. 1981. Monoclonal antibodies to
herpes simplex virus type 1 proteins, including the immediate-early pro-
tein ICP 4. Infect. Immun. 34:684–692.
40. Simbulan-Rosenthal CM, Rosenthal DS, Iyer S, Boulares AH, Smulson
ME. 1998. Transient poly(ADP-ribosyl)ation of nuclear proteins and role
of poly(ADP-ribose) polymerase in the early stages of apoptosis. J. Biol.
Activation of PARP by HSV-1
August 2012 Volume 86 Number 15 jvi.asm.org 8267
41. Smith MC, Boutell C, Davido DJ. 2011. HSV-1 ICP0: paving the way for Download full-text
viral replication. Future Virol. 6:421–429.
42. Sodhi RK, Singh N, Jaggi AS. 2010. Poly(ADP-ribose) polymerase-1
(PARP-1) and its therapeutic implications. Vascul. Pharmacol. 53:77–87.
assessment for suspension cultures of mammalian cells. Biotechnol. Bio-
44. Vastag L, Koyuncu E, Grady SL, Shenk TE, Rabinowitz JD. 2011. Divergent
45. Wang Y, et al. 2011. Poly(ADP-Ribose) (PAR) binding to apoptosis-
inducing factor is critical for PAR polymerase-1-dependent cell death
(Parthanatos). Sci. Signal. 4:ra20. doi:10.1126/scisignal.2000902.
46. Weitzman M, Weller S. 2011. Interactions between HSV-1 and the DNA
Press, Norfolk, VA.
47. Whatcott CJ, Meyer-Ficca ML, Meyer RG, Jacobson MK. 2009. A
specific isoform of poly(ADP-ribose) glycohydrolase is targeted to the
mitochondrial matrix by a N-terminal mitochondrial targeting sequence.
Exp. Cell Res. 315:3477–3485.
48. Ying W, Sevigny MB, Chen Y, Swanson RA. 2001. Poly(ADP-ribose)
glycohydrolase mediates oxidative and excitotoxic neuronal death. Proc.
Natl. Acad. Sci. U. S. A. 98:12227–12232.
49. Yuan J, Bennett BD, Rabinowitz JD. 2008. Kinetic flux profiling for
quantitation of cellular metabolic fluxes. Nat. Protoc. 3:1328–1340.
50. Zhou Y, Feng X, Koh DW. 2011. Activation of cell death mediated by
apoptosis-inducing factor due to the absence of poly(ADP-ribose) glyco-
hydrolase. Biochemistry 50:2850–2859.
Grady et al.
jvi.asm.org Journal of Virology